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Uracil DNA Glycosylase: Insights from a Master Catalyst
Archives of Biochemistry and Biophysics Vol. 396, No. 1, December 1, pp. 1–9, 2001 doi:10.1006/abbi.2001.2605, available online at http://www.idealibrary.com on
MINIREVIEW Uracil DNA Glycosylase: Insights from a Master Catalyst 1 James T. Stivers 2 and Alexander C. Drohat Department of Pharmacology and Molecular Sciences, The Johns Hopkins University School of Medicine, 725 North Wolfe Street, Baltimore, Maryland 21205-2185
Received September 12, 2001
The recognition and removal of damaged bases in the genome is the province of a highly specialized assemblage of enzymes known as DNA glycosylases. In recent years, structural and mechanistic studies have rapidly moved forward such that in some cases, the high-resolution structures of all stable complexes along the reaction pathway are available. In parallel, advances in isotopic labeling of DNA have allowed the determination of a transition state structure of a DNA repair glycosylase using kinetic isotope effect methods. The use of stable substrate analogs and fluorescent probes have provided methods for real time measurement of the critical step of damaged base flipping. Taken together, these synergistic structural and chemical approaches have elevated our understanding of DNA repair enzymology to the level previously attained in only a select few enzymatic systems. This review summarizes recent studies of the paradigm enzyme, uracil DNA glycosylase, and discusses future areas for investigation in this field. © 2001 Elsevier Science Key Words: DNA glycosylase; base flipping; inhibition; chemical rescue; transition state structure; enzyme mechanism.
DNA has evolved to be the genetic storage material because of its exceptional, but not infinite chemical stability. Nevertheless, genomic DNA is often viewed as a frail target that suffers the onslaught of a variety of chemical insults resulting in deamination, oxidation, alkylation, or other forms of chemical damage to the natural bases (1). Such damage, although infrequent, 1 The work in the author’s laboratory is supported by National Institutes of Health Grant GM46835. 2 To whom correspondence and reprint requests should be addressed. Fax: (410) 955-3023. E-mail: [email protected].
0003-9861/01 $35.00 © 2001 Elsevier Science All rights reserved.
is intolerable with respect to the preservation of the coding potential of the genome, and exerts the selection pressure for evolution of an elaborate cellular DNA repair machinery. Paradoxically, the intrinsic stability of DNA makes the breaking of its chemical bonds during DNA repair an energetically challenging feat, and therefore requires the action of powerful enzyme catalysts to efficiently recognize and remove damaged bases through a pathway known as base excision repair (Fig. 1). The first step in this repair pathway is the hydrolytic cleavage of the N-glycosidic bond of the damaged base by a damage specific DNA glycosylase, which is the focus of this review. DNA glycosylases face two fundamental obstacles in the initiation of this repair process. First, these enzymes must solve the formidable challenge of locating damaged bases in the vast genome before these potentially mutagenic sites are fixed by DNA replication. This critical time frame provides the driving force for selection of efficient damage search and detection mechanisms. Such mechanisms may involve three-dimensional or one-dimensional search processes (2), or even processive scanning mechanisms in which unusual features of the damage site are detected by the enzyme (3). A number of high resolution crystal structures of DNA repair glycosylases bound to DNA have appeared in recent years that have revealed several unifying principles of the damage recognition process (4 –10). All monofunctional DNA repair glycosylases studied to date operate by a base flipping mechanism in which the damaged base is extruded from the DNA base stack into the active site pocket of the enzyme, which is highly complementary to the shape and electronic features of the base. The remarkable conservation of the base flipping mechanism attests to the importance of gaining access to the functional groups on the base and sugar, which provides the binding inter1
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STIVERS AND DROHAT TABLE I
Stabilities of Representative Deoxynucleosides at pH 7 and pH 3 and Estimated Catalytic Powers of Several DNA Glycosylases a
Nucleoside
t 1/2 (pH 7, 37°C)
t 1/2 (pH 3, 37°C)
DNA context
Specific DNA glycosylase
Catalytic power b (pH 7, 37°C)
T dU dA N 3-Me-dA
109 years 23 years 3.9 years 36 min
109 years 23 years 6.8 h 3 min
T/G U/A, U/G A/8-oxo-dG N 3-Me-dA/T
TDG UDG MutY TAG
7.5 ⫻ 10 7 2 ⫻ 10 12 4 ⫻ 10 7 ⱖ6
a This is not a comprehensive survey of all types of damaged bases, but does cover the entire range of stabilities that are found in nature. The half-lives for the nucleosides were calculated from the rate constants and activation parameters in references (20, 34, 35). b The catalytic power is defined as the half-life for the nonenzymatic reaction divided by that for the enzymatic reaction and is a measure of how much the enzyme lowers the activation barrier as compared to the nonenzymatic reaction. The enzymatic rates were obtained from single turnover measurements, such that slow product release rates do not affect the comparison. The enzymatic rates were obtained from the following studies: thymine DNA glycosylase (TDG) (36); uracil DNA glycoyslase (37); adenine mismatch-specific glycosylase (MutY) (38); and 3-methyladenine DNA glycosylase I from E. coli (TAG) (39).
actions necessary for lowering the activation barrier of the reaction. The stability of the glycosidic bond in pyrimidine, purine and alkylated purine deoxynucleosides varies greatly (Table I), which places differing catalytic demands on each of these enzymes. However, there are two fundamental chemical problems that each enzyme must solve to varying degrees. The first involves providing a mechanism for stabilization of the increased electron density that accumulates on the base during
bond cleavage. The second is balancing the degree of bond formation to the water nucleophile and bond breakage to the leaving group base in the transition state. The latter issue is a key mechanistic point because if the base departs completely before the water nucleophile attacks, the enzyme active site architecture must be designed to stabilize a highly unstable positively charged oxacarbenium ion intermediate (Fig. 1). In contrast, a transition state involving a large amount of bond formation to the water nucleophile
FIG. 1. Pathway for DNA base excision repair. Cellular DNA is damaged by a variety of exogenous and endogenous agents, resulting in alterations in the covalent structure of the natural bases (B*). The first step in repair is the hydrolytic cleavage of the N-glycosidic bond by a damage specific DNA glycosylase. The chemical mechanism of bond cleavage may be associative (upper pathway), or dissociative (lower pathway) involving a discrete positively charged oxacarbenium ion intermediate (see text). The repair process is completed by the action of an apyrimidinic endonuclease (AP endo), a repair polymerase (pol ), and DNA ligase that collectively remove the abasic nucleotide and reinsert the correct dNTP.
DNA GLYCOSYLASE MECHANISMS
3
requires activation of the anomeric carbon atom, which is weakly electrophilic, as well as the attacking water, which is a weak nucleophile. Of course, these issues are not new, and simply illustrate another example of the long-standing question of whether enzymes exert their power by stabilizing otherwise unstable intermediates (lower pathway, Fig. 1), or use catalytic strategies to lower the energy of concerted transition states (upper pathway, Fig. 1) (11). As will be discussed below, the electronic features of the base and the deoxyribose sugar in the transition state or high energy intermediate are likely to play a key role in the rational design of inhibitors of these enzymes. The intent of this review is to summarize some of the chemical approaches that have been used in recent years to elucidate the site recognition and catalytic mechanisms of the enzyme, uracil DNA glycosylase (UDG). 3 An emphasis is placed on UDG because the structural and mechanistic studies have advanced most rapidly in this system, and should therefore serve as a paradigm for similar studies of other glycosylases. DAMAGE RECOGNITION AND URACIL FLIPPING
Uracil in DNA may arise from either the spontaneous deamination of cytosine, resulting in U:G mismatches, or from the misincorporation of dUTP instead of TTP during DNA replication, which results in U:A base pairs (12). Such processes occur with alarming frequencies in normal cells (10 2 to 10 3 per cell per day), requiring constant surveillance and removal. The selective removal of a uracil base in the context of U:G or U:A base pairs, without also removing the structurally related base T, is one of the most difficult specific recognition problems in DNA repair. The entire process of damage site recognition is still not very well understood, but may be conceptualized as a three step process: (1) a three-dimensional or one dimensional search process in which the site is located in a sea of normal DNA base pairs, (2) the formation of largely nonspecific interactions with the site prior to base flipping, and (3) the formation of specific interactions with the flipped out nucleotide. The high-resolution crystal structures of UDG bound to specific DNA have contributed greatly to an understanding of the final step of extrahelical recognition (7, 10, 13). However, an understanding of the earlier steps requires either extrapolation backwards in time starting from the crystal models, or the use of other approaches that are better suited for understanding transient complexes. The specific recognition process of UDG is discussed within the context of this three-step model. 3 Abbreviations used: UDG, uracil DNA glycosylase; wt, wild-type; P, 2-aminopurine; MUG, mismatch uracil DNA glycoylase; TDG, thymine DNA glycosylase.
FIG. 2. (A) Structural model of the active site interactions of human UDG bound to duplex DNA containing the C-glycoside analogue 2⬘-deoxypseudouridine (dU) (PDB code 1 emh). Various residues proposed to be involved in chemical catalysis and glycosidic bond strain are indicted (see text); the residue numbering corresponds to the E. coli enzyme. (B) Hypothetical tautomerization reaction of dU that could account for the highly unusual distorted tetrahedral geometry of C1 observed in the crystal structure in A (see text).
Structure of the extrahelical complex. The crystal structure of human UDG bound to DNA containing the stable C-glycoside analog of deoxyuridine, deoxypseudouridine (dU, Fig. 2A, 2B), reveals the salient features of the site specific complex in which the uracil base is extrahelical. Aside from the unusual glycosidic bond angle seen in this structure (discussed later), this complex shares several features observed in other glycosylase complexes with extrahelical bases. First, UDG is found to use hydrogen bond donors (Ser88, Ser189) to clamp and pinch the phosphodiester backbone of the DNA on the same strand that contains the uracil base, leading to localized DNA bending (⬃45°). Another conserved aspect of base flipping seen in this structure is that a bulky hydrophobic amino acid side chain (Leu191, not shown for clarity) is found to project into the minor groove of the DNA duplex, filling the void left by the departed uracil base. The role of this enzyme group may be to actively push the damaged base from
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STIVERS AND DROHAT
FIG. 3. Conceptual model for uracil base flipping by UDG. The overall process of base flipping may be viewed as being comprised of two components: an unfavorable duplex destabilization term (⌬G D), and a favorable stabilizing free energy (⌬G S) arising from all the interactions gained by flipping the base into the active site. Although poorly characterized, duplex destabilization is thought to arise in part from phosphodiester compression that occurs when UDG binds nonspecifically to DNA. The favorable free energy term arises in part from specific hydrogen bonding and stacking interactions with the uracil base, and intercalation of Leu191 into the DNA base stack to hinder reinsertion of the departed uracil.
the DNA stack, or alternatively, simply to hinder its reinsertion once the extrahelical state is attained by other means (see below). Finally, several completely conserved enzyme side chains form specific hydrogen bonds and pull the uracil base into the binding pocket: the O4 and H3 Watson-Crick acceptor and donor pair interact with the side chain atoms of Asn123, and uracil O2 forms hydrogen bonds to N 2H of His187, and the backbone amide of Gln63 (not shown). To venerate these observed interactions, a “pinch–push–pull” mechanism for base flipping has been proposed (6). Another key realization from this structure is that the specificity for uracil, as opposed to thymine, likely arises from unfavorable steric interactions of the 5-methyl group of T with Tyr66, which forms a rigid wall on one side of the binding pocket. Consistent with this hypothesis, removal of Tyr66 converts UDG into a thymine DNA glycosylase (14). Detecting transient complexes on the base flipping pathway. How do the initial interactions with the damage site and undamaged DNA lead to uracil flipping? Currently, there is no structure of a DNA glycosylase bound to undamaged DNA, or alternatively, to a damaged DNA site in which the base is not extrahelical. Thus, an entirely satisfying answer to this question is not yet in hand— but a trail for pursuit does exist. As suggested by both structural and biophysical studies, the earliest interaction of UDG with DNA probably involves local compression of the DNA phos-
phodiester backbone, perhaps leading to torsional stress in the DNA duplex. Upon encounter with the uracil site, the stress is presumably relieved by flipping the entire deoxyuridine nucleotide from the helical stack into the active site pocket of the enzyme (Fig. 3). The evidence for such a strain and release mechanism is indirect, but compelling. First, UDG has been shown to bind weakly to undamaged DNA (K D ⫽ 2 to 5 M), and much more tightly to DNA that contains the stable substrate analog 2⬘--fluoro-2⬘-deoxyuridine (2⬘FdU, K D ⬃25 nM). Raman spectroscopy and chemical probe methods have begun to characterize some features of the nonspecific and specific interactions with DNA. One spectral feature of the nonspecific complex was Raman hypochromism (analogous to UV hypochromism), arising from increased base stacking interactions in the DNA upon UDG binding (15). This spectral signature was also observed in the specific complex with substrate analog DNA, suggesting shared structural features in both complexes. Since Raman is a very sensitive probe for – interactions, the increase in stacking could be reporting indirectly on DNA distortion arising from enzyme interactions with the phosphodiester backbone. In a complementary approach, permanganate oxidation sensitivity experiments have been used to probe the change in accessibility of thymine bases upon binding of UDG to an undamaged 24-mer duplex DNA (Y. L. Jiang and J. T. Stivers, unpublished). These experiments clearly showed that UDG binding increased the oxidation sensitivity of specific T residues, to a level approaching that of the same sequence in a ssDNA context. These results lead us to hypothesize that UDG induces a subtle localized distortion in the duplex upon nonspecific binding that can destabilize base pairs some distance from the site of interaction. Although neither the Raman or chemical probe results can unambiguously establish that duplex destabilization is occurring, both methods reveal changes in the duplex structure that are evocative of such a mechanism. Another useful approach to study the steps leading to base flipping has been to use the fluorescent nucleotide probe 2-aminopurine (2-AP) in combination with the stable dU substrate analogue, 2⬘-fluoro-2⬘-deoxyuridine (2⬘-FdU) (16). 2-Aminopurine is an adenine analogue that has the useful properties of retaining base pair complementarity with T (although one hydrogen bond is moved to the minor groove of the duplex) and a fluorescence quantum yield that is very sensitive to stacking interactions with other bases in the duplex (17). Accordingly, when 2-AP ( ex ⫽ 315 nm, em ⫽ 370) is placed adjacent to 2⬘-FdU in duplex DNA, there is a large decrease in the stacking interaction when the uracil is flipped into the UDG active site, resulting in an eightfold increase in fluorescence. Such an approach provided the basis for a stopped-flow ki-
DNA GLYCOSYLASE MECHANISMS
netic analysis that revealed a two-step mechanism for damage site recognition and uracil flipping by UDG. In the first step, UDG forms an unstable complex in a diffusion controlled association reaction (K D ⬃ 2 M). It is noteworthy that this complex is kinetically and thermodynamically indistinguishable from the complex UDG forms with undamaged DNA, suggesting that the initial stage in site recognition is the formation of nonspecific interactions with the duplex. Once the weak complex forms, it may then decay backwards to form free reactants (k diss ⬃ 700 s ⫺1) or forward to form the specific extrahelical conformation (k flip ⬃ 1200 s ⫺1). The extremely rapid rate of dissociation from the weak complex is not consistent with a highly processive one-dimensional search mechanism, but is consistent with previous processivity measurements of eUDG (in the presence of physiological concentrations of salt), which indicated the enzyme did not linearly diffuse long distances on the DNA before dissociation (x). Another interesting finding from the fluorescence studies was that the internal equilibrium constant for base flipping on the enzyme (K flip ⫽ 12 to 33) was similar for duplex DNA containing 2⬘-FdU/A and 2⬘FdU/G base pairs, as well as single stranded 2⬘-FUcontaining DNA. This key observation provides additional support for a duplex destabilization step prior to base flipping, because it is difficult to rationalize the similar base flipping rates and equilibria for duplex and single stranded DNA unless the duplex DNA was destabilized prior to the flipping step. Stopped-flow tryptophan fluorescence measurements also indicate that an induced fit conformational change in UDG accompanies the process of uracil flipping. The conformational change was also detected by Raman spectroscopy when UDG binds to DNA containing 2⬘-FdU, but not undamaged DNA, indicating that the conformation of the enzyme is altered by the specific interactions that are formed as a result of base flipping (15, 16). Such a conformational change, which occurs concomitantly with base flipping, argues strongly for direct involvement of the enzyme in the formation of the extrahelical state, rather than passive trapping of a spontaneously formed extrahelical base (18). Manipulating base flipping using mutagenesis and chemical approaches. Two approaches to test our understanding of a biological reaction are to generate mutations, or alternatively, synthesize substrate analogs that remove functional groups that are hypothesized to play a role in the process. Conversely, the generation of substrate analogs or small molecules that “rescue” the damaging effects of enzyme mutations can provide valuable insights into the role of the enzyme functional group in the mechanism. One novel example of the chemical rescue approach was recently employed
5
FIG. 4. A pyrene nucleotide to promotes uracil flipping. (A) The bulky pyrene nucleotide fills the entire space usually occupied by a normal base pair. Thus substitution of pyrene (Y) for the A opposite to U (i.e., U/A 3 U/Y) would be expected to push the uracil from the base stack and facilitate specific DNA binding by diminishing the free energy penalty for duplex destabilization prior to uracil flipping (see Fig. 3). (B) Schematic depiction of the mechanism of pyrene (hashed symbol) assisted flipping (19).
to probe the pushing role of Leu191 in uracil flipping by UDG (19). Deletion of Leu191 results in a 10 to 625-fold decrease in k cat/K m using duplex DNA substrates containing a U:A base pair, presumably because UDG can no longer stabilize the extrahelical uracil in the active site. This interpretation was tested by substituting a pyrene nucleotide analogue for adenine opposite to the uracil (Fig. 4A). This rational design approach was based on the idea that the bulky pyrene “base,” which fills the entire space normally occupied by the normal U:A base pair (Fig. 4B), might serve as a mechanical wedge to either force the uracil from the DNA base stack in the free DNA (pushing) or hinder its reinsertion once it is expelled in the UDG complex (plugging). We predicted that Y would selectively rescue the L191A mutant of UDG that is defective in promoting the extrahelical base and would have a lesser effect on wtUDG and other mutants that were not defective in the base flipping step. Consistent with these expectations, pyrene opposite to U (a U:Y pair) selectively and completely rescued the impaired catalytic and binding activities of the L191A pushing mutant of UDG, with only a small salutary effect on mutants that were not deficient in base flipping. A model was proposed in which pyrene diminishes the energy penalty for flipping the base from the duplex, and stabilizes the extrahelical state. Pyrene rescue should serve as a useful tool to diagnose the functional roles of other residues involved in promoting the extrahelical base.
6
STIVERS AND DROHAT
CATALYTIC STRATEGIES
Glycosidic bond hydrolysis and the intrinsic chemical properties of deoxynucleotides. Before considering the nature of enzymatic catalysis it is always useful to identify the character of the chemical problem through consideration of the nonenzymatic reaction. The halflives and pH dependences of the nonenzymatic glycosidic bond cleavage reactions of pyrimidines, purines, and alkylated purine deoxynucleosides illustrate the point that the various DNA glycosylases specific for these base types face unique catalytic problems arising from the different intrinsic chemical properties of the bases (Table I). Pyrimidine-specific enzymes such as uracil DNA glycosylase (UDG) and the U:G and T:G mismatch glycosylases, MUG and TDG, face the most difficult leaving group activation problem because U and T cannot be readily protonated (pK a ⬍ ⫺3.4) (20). Purine bases are quite stable to hydrolysis at neutral pH (t 1/ 2 ⬃ 4 yrs). However, these bases are more easily protonated (pK aN1 ⫽ 3.5, pK aN7 ⬍ 1) (21), which provides a powerful mechanism for enzymatic activation of these bases by enzymes such as MutY (Table I). Finally, alkylated purine bases, and especially 3-MeA, are quite labile even at neutral pH (t 1/ 2 ⫽ 36 min), and may not require any activation at all by the alkyl purine specific enzymes such as 3-methyladenine DNA glycosylase I from Escherichia coli (TAG, Table I). Accordingly, the catalytic power of these enzymes 4 varies wildly from as little as 10-fold for TAG, to as much as 10 12-fold for UDG at pH 7 and 37°C. Of course, leaving group activation is not the only catalytic strategy that may be employed. If the enzymatic reaction pathway follows a highly dissociative mechanism in which a positively charged oxacarbenium ion is generated (Fig. 1), catalysis would be enhanced by any strategy that leads to stabilization of this cationic species. Such strategies could involve electrostatic mechanisms, or the use of binding energy to distort the sugar into a conformation that favors stabilizing hyperconjugative effects from the 2⬘ hydrogens in the dissociative transition state or intermediate (22). If an associative mechanism involving significant bond formation to the water nucleophile is followed, then such strategies as general base catalysis of water attack and polarization of the anomeric carbon would become useful. The following discussion of UDG uses the available structural and mechanistic information to evaluate the significance of these potential catalytic strategies. It is likely that related strategies will be used by other DNA glycosylases. 4 In this review, the catalytic power of an enzyme is defined as the single-turnover rate of the enzyme catalyzed reaction divided by the rate of the uncatalyzed reaction. Although k cat is often used for this comparison, this rate constant is less meaningful in the case of DNA glycosylases because the product release step is rate limiting.
Evaluating ground state strain and stereoelectronic effects with UDG. UDG is the most powerful DNA glycosylase yet identified. The enzyme provides a rate acceleration of 10 12-fold over the uncatalyzed hydrolysis reaction at 25°C, placing it in the upper echelon of enzyme catalysts. The study of UDG has been surprisingly amenable to a wide range of biophysical approaches. Extensive crystallographic studies on the human and E. coli enzymes have yielded structures of each complex along the reaction pathway (6, 7, 10, 23). In addition, heteronuclear NMR, Raman spectroscopic and kinetic isotope effect studies have provided unique perspectives into each reaction complex, including the transition state. One of the least understood and most interesting aspects of UDG catalysis is the role of ground state strain and stereoelectronic effects. These questions were propelled into serious consideration by the provocative and compelling crystal structure of human UDG in complex with DNA containing the stable Cglycoside substrate analog 2⬘-deoxypseudouridine (⌿dU, Fig. 2A). This structure indicates that the normal trigonal planar C1 position of ⌿dU is bent toward a tetrahedral geometry, presumably through steric interactions with a conserved phenylalanine in the active site, and the 3⬘ and 5⬘ serine phosphodiester clamps. It was argued that such an unusual conformation, if affected in the natural substrate, would lead to groundstate bond strain and enhanced electron orbital overlap between the lone pair electrons on O4⬘ and the -orbitals of the uracil ring in the transition state, thereby solving a problem described as the “orthogonal lone pair paradox” (7). 5 The interpretation of this structure requires contemplation of two separate questions. First, is ⌿dU-DNA a faithful substrate mimic? And second, is such a destabilization mechanism energetically reasonable and, more importantly, catalytic? The available evidence indicates that ⌿dU is not the best available substrate mimic for UDG. DNA containing ⌿dU binds 20-fold more weakly than the same DNA sequence containing the alternative slow substrate and competitive inhibitor 2⬘-FdU (Y. L. Jiang and J. T. Stivers, unpublished), which has a K D similar to the K m of the natural 2⬘-dU-containing substrate (16). It is also known from Raman spectroscopy studies that 2⬘-FdU in DNA shows a large 34 cm ⫺1 downshift in 5
The term paradox was used to describe the orthogonal arrangement of the * orbital of the anomeric carbon and the orbital of the C2–O2 bond of the uracil ring in the normal anti-conformation of deoxyuridine (see Fig. 5 of Ref. 7). There is a long standing debate on the energetic importance of such stereoelectronic effects (33). In the present case, there is experimental evidence from Raman spectroscopy for N ␦⫹ ⫽ C-O ␦⫺ zwitterionic character at N1–C2 ⫽ O in the ground state. The presence of such a resonance form suggests that alignment of orbitals would not be required for efficient leaving group expulsion in the transition state.
DNA GLYCOSYLASE MECHANISMS
the carbonyl stretching frequencies of the uracil ring upon binding to UDG, indicating strong hydrogen bonding to active site groups (Fig. 2A), whereas ⌿dU containing DNA does not show any significant polarization of these carbonyl groups (J. Dong and P. R. Carey, unpublished). An additional concern, that is entirely unresolved, is that the observed structure for ⌿dU in the crystal could arise not from mechanical strain, but from a tautomerization reaction, in which the H3 proton is transferred to C1 (Fig. 2B). Indeed, the observed crystal conformation and the computationally optimized structure of the C1-H tautomer of deoxypseudouridine are very similar, and probably beyond the resolution of the diffraction data (1.8 Å). Although QM calculations indicate that this tautomerization reaction is thermodynamically unfavorable in solution by about 20 kcal/ mol, due to the loss of aromaticity in the ring (J. Dong and P. R. Carey, unpublished), the active site environment of UDG could kinetically facilitate this reaction through the use of the strong H-bond with His187 (Fig. 2, see also below). The proton could become trapped at C1 if no basic group is available to accept the proton in the tetrahedral tautomer (no such group is seen in the crystal structure). Although this thermodynamic barrier is large, there are many enzymes in Nature that catalyze similar unfavorable tautomerization reactions using electrophilic mechanisms (24). A perplexing observation is that no unusual changes in the normal modes of the uracil ring were observed in solution Raman studies of 2⬘-FdU bound to UDG, as would be expected from such bond distortion. This is intriguing because 2⬘-FdU cannot undergo the same tautomerization reaction as ⌿dU. In summary, the questions raised by this structure have large implications for the role of strain in enzyme catalysis, and more specifically, in the design of tight binding inhibitors of UDG. A rational viewpoint for the enzymologist at this juncture is to await the results of further structural or spectroscopic studies that will undoubtedly clarify the role of ground state strain in this system. If such a mechanism is found to prevail, it will be fascinating to dissect the mechanical forces that give rise to such a remarkable effect. Nature of the transition state. Significant advances have been made in understanding the electronic and structural features of the transition state of the UDG reaction using spectroscopic and kinetic isotope effect methodologies. The first hint at the electronic character of the transition state came from 2D 13C heteronuclear NMR studies, in which the ionization state of the uracil base in the ternary product complex with abasic DNA was determined (25). Quite surprisingly, the uracil was found to have a dramatically lowered N1 pK a, such that the uracil anion was the sole bound
7
species at neutral pH (pK aN1 ⫽ 6.4). The presence of the uracil anion was later confirmed in Raman studies (15). Taken together, these observations indicated that the enzyme stabilized the uracil anion leaving group by 5 kcal/mol in the product complex as compared to free uracil in solution (pK aN1 ⫽ 9.8)— but how? NMR spectroscopy also illuminated the answer to this energetic problem. Inspection of the 1D proton NMR spectrum of the UDG complex with uracil and abasic DNA revealed a downfield shifted proton (␦ ⫽ 15.7 ppm), which was unambiguously assigned to a short (⬍2.7 Å) hydrogen bond between the O2 anion of uracil and the N 2 H of the conserved His187 (26). Histidine 187 was previously shown by NMR to be neutral in the pH range 4.5–10, which, in addition to the other NMR evidence, established the neutral histidine-uracil anion interaction depicted in Fig. 5. This provided yet another example in which evolution has selected for a short strong hydrogen bond from a neutral histidine to stabilize a developing negative charge on an enolic oxygen atom (27). Thus, part of the catalytic story of UDG involves leaving group stabilization by hydrogen bonding, but not proton transfer. An inference from these NMR studies was that the uracil anion might also exist as an intermediate in a stepwise mechanism for glycosidic bond cleavage. The direct determination of the transition state structure for the UDG catalyzed reaction was obtained from kinetic isotope effect measurements. This approach required the development of methods for the enzymatic synthesis of oligonucleotide substrates containing stable and radioactive isotope labels at a single uracil residue (28, 29). As discussed in two recent and excellent reviews by Schramm and Berti (30, 31), kinetic isotope effects (KIE ⫽ k light/k heavy) measure the changes in vibrational frequencies for heavy and light isotopes incorporated at specific atomic sites of a reactant as it is transformed from its reactant state to the transition state geometry, and thus provide information on bond lengths and geometries in the transition state. The experimental KIEs for UDG unambiguously indicated a dissociative transition state with substantial oxacarbenium ion character, and confirmed the proposal that a short-lived oxacarbenium ion-uracil anion intermediate is formed on the enzyme (Fig. 5). One remarkable feature of this intermediate was that the sugar adopted an unusual 3⬘-exo pucker, which allows maximal stabilizing hyperconjugative effects from the 2⬘  protons to the electron deficient anomeric carbon. This sugar conformation in the intermediate was strikingly similar to that observed in the crystal structure of the Michaelis complex, suggesting that UDG uses binding energy in the ground state to preorganize the substrate into this reactive conformation. We concluded that several features of the active site contributed to the increased stability of the intermediate as
8
STIVERS AND DROHAT
FIG. 5. Interactions leading to stabilization of the oxacarbenium ion-uracil anion intermediate in the UDG active site. The electrostatic potentials for the dU reactant and the oxacarbenium ion intermediate are projected onto the van der Waals surfaces for each molecule. The structures were derived using the geometric information derived from KIE measurements (28), and the crystal coordinates of the product complex with the uracil anion and abasic DNA (PDB code 1 ssp). For clarity, His187 is not shown in the depiction of the electrostatic potentials. Stabilization of the ionic intermediate is thought to arise from the enforcement of stabilizing hyperconjugative effects in the reactant and transition states using the serine phosphodiester interactions, and (ii) electrostatic stabilization of the oxacarbenium ion by the anionic uracil leaving group and Asp64.
compared to solution: (i) the enforcement of stabilizing hyperconjugative effects in the reactant and transition states using the serine phosphodiester interactions, and (ii) electrostatic stabilization of the oxacarbenium ion by the anionic uracil leaving group and Asp64 (Fig. 5). Substrate binding energy and catalysis. Although enzymes are sometimes viewed as elaborate scaffolds for carrying chemical catalysts, the intricate interplay between noncovalent interactions with the substrate that are distant from the site of chemistry can play an important role in catalysis. Such interactions are critical for obtaining proper substrate positioning with respect to catalytic groups, driving conformational changes in the substrate that enhance its reactivity, and in induced fit specificity mechanisms. One approach to investigate these effects with UDG was to systematically dismantle the optimal substrate, Ap ⫹1UpA ⫺1pA ⫺2, by replacing the nucleotides at the ⫹1, ⫺1, or ⫺2 positions with a tetrahydrofuran abasic site analogue, 3-hydroxypropyl phosphodiester spacer, a phosphate monoester, or a hydroxyl group (32). Thus, base, phosphodiester, and deoxyribose interactions were removed in a controlled fashion, and the damaging effects on catalysis could be ascertained. One key finding in this extensive study was that the minimal substrate for UDG is deoxyuridine (dU). Although four billion-fold less reactive than the optimal substrate, cleavage of the glycosidic bond of dU was still enhanced by 4 ⫻ 10 7-fold over the spontaneous reaction in the absence of UDG. A second significant finding was the tremendous importance of the ⫺2 nucleotide in driving
optimal positioning of the substrate. The net catalytic benefit of adding a ⫺2 adenine nucleotide to ApUpA (i.e., ApUpA 3 ApUpApA) was 270,000-fold (⫺7.5 kcal/ mol). The crystal structure suggests a structural basis for this large effect because the pro-Sp nonbridging oxygen atom of the ⫺2 phosphodiester is hydrogen bonded to the backbone amide group of the catalytic electrophile His187, which is in the same loop as Leu191 (7). Thus the ⫺2 nucleotide likely serves as a key handle to drive the active site into the productive closed conformation, allowing formation of the strong hydrogen bond between uracil O2 and His187, optimizing the serine-phosphodiester interactions at the ⫹1 and ⫺1 positions, and positioning Leu191 in the minor groove. FUTURE PROSPECTS
There still remain many interesting questions to be investigated with respect to the mechanism of damage site recognition and base flipping by UDG and other repair glycosylases. One exciting area for exploration would be the use of single molecule fluorescence methods to visualize the process of damage site location one molecule at a time. New approaches also need to be used to understand the structure and dynamics of the nonspecific complex between UDG and undamaged DNA. It is likely that NMR will find great utility in this respect, as it may be possible to determine the structure of duplex DNA in such a complex, and to investigate the dynamics by measuring imino proton exchange rates or performing 13C or 15N relaxation mea-
DNA GLYCOSYLASE MECHANISMS
surements. These NMR dynamic studies could be augmented with time-resolved fluorescence measurements of the dynamic behavior of damaged DNA and its complex with the enzyme. Such investigations have already been performed on the complex of UDG with the abasic product, revealing that the enzyme restricts the conformational dynamics of the duplex compared to the free DNA (40). Finally, it would be of great utility to develop selective small molecule inhibitors of DNA glycosylases and other enzymes involved in DNA base excision repair. Cell permeable inhibitors could be used to better probe the biological role of this pathway under various stress conditions such as those used in cancer chemotherapy treatment. The pursuit of these and other goals will continue to make DNA repair an exciting field into the foreseeable future. REFERENCES 1. Friedberg, E. C. (1985) in DNA Repair, W. H. Freeman, New York, NY. 2. Berg, O. G., Winter, R. B., and von Hippel, P. H. (1981) Biochemistry 20, 6929 – 6948. 3. Verdine, G. L., and Bruner, S. D. (1997) Chem. Biol. 4, 329 –334. 4. Lau, A. Y., Scharer, O. D., Samson, L., Verdine, G. L., and Ellenberger, T. (1998) Cell 95, 249 –258. 5. Lau, A. Y., Wyatt, M. D., Glassner, B. J., Samson, L. D., and Ellenberger, T. (2000) Proc. Natl. Acad. Sci. USA 97, 13573– 13578. 6. Parikh, S. S., Mol, C. D., Slupphaug, G., Bharati, S., Krokan, H. E., and Tainer, J. A. (1998) EMBO. J. 17, 5214 –5226. 7. Parikh, S. S., Walcher, G., Jones, G. D., Slupphaug, G., Krokan, H. E., Blackburn, G. M., and Tainer, J. A. (2000) Proc. Natl. Acad. Sci. USA 97, 5083–5088. 8. Bruner, S. D., Norman, D. P., and Verdine, G. L. (2000) Nature 403, 859 – 866. 9. Barrett, T. E., Scharer, O. D., Savva, R., Brown, T., Jiricny, J., Verdine, G. L., and Pearl, L. H. (1999) EMBO J. 18, 6599 – 6609. 10. Werner, R. M., Jiang, Y. L., Gordley, R. G., Jagadeesh, G. J., Ladner, J. E., Xiao, G., Tordova, M., Gilliland, G. L., and Stivers, J. T. (2000) Biochemistry 39, 12585–12594. 11. Jencks, W. P. (1980) Accounts Chem. Res. 13, 161–169. 12. Mosbaugh, D. W., and Bennett, S. E. (1994) Prog. Nucleic Acid Res. Mol. Biol. 48, 315–370. 13. Parikh, S. S., Mol, C. D., and Tainer, J. A. (1997) Structure 5, 1543–1550.
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14. Kavli, B., Slupphaug, G., Mol, C. D., Arvai, A. S., Peterson, S. B., Tainer, J. A., and Krokan, H. E. (1996) EMBO J. 15, 3442–3447. 15. Dong, J., Drohat, A. C., Stivers, J. T., Pankiewicz, K. W., and Carey, P. R. (2000) Biochemistry 39. 16. Stivers, J. T., Pankiewicz, K. W., and Watanabe, K. A. (1999) Biochemistry 38, 952–963. 17. Stivers, J. T. (1998) Nucleic Acids Res. 26, 3837–3844. 18. Cheng, X., and Blumenthal, R. M. (1996) Structure 4, 639 – 645. 19. Jiang, Y. L., Kwon, K., and Stivers, J. T. (2001) J. Biol. Chem., submitted. 20. Shapiro, R., and Danzig, M. (1972) Biochemistry 11, 23–29. 21. Blackburn, G. M., and Gait, M. J. (1996) in Nucleic Acids in Chemistry and Biology, Oxford Univ. Press, Oxford. 22. Sunko, D. E., Szele, I., and Hehre, W. J. (1977) J. Am. Chem. Soc. 99, 5000 –5005. 23. Xiao, G., Tordova, M., Jagadeesh, J., Drohat, A. C., Stivers, J. T., and Gilliland, G. L. (1999) Proteins 35, 13–24. 24. Gerlt, J. A., and Gassman, P. G. (1993) Biochemistry 32, 11943– 11952. 25. Drohat, A. C., and Stivers, J. T. (2000) J. Am. Chem. Soc. 1840 – 1841. 26. Drohat, A. C., and Stivers, J. T. (2000) Biochemistry 39, 11865– 11875. 27. Cleland, W. W., Frey, P. A., and Gerlt, J. A. (1998) J. Biol. Chem. 273, 25529 –25532. 28. Werner, R. M., and Stivers, J. T. (2000) Biochemistry 39. 29. Gilles, A.-M., Cristea, I., Palibroda, N., Hilden, I., Jensen, K. F., Sarfati, R. S., Namane, A., Ughetto-Monfrin, J., and Barzu, O. (1995) Anal. Biochem. 232, 197–203. 30. Schramm, V. L. (1999) Methods Enzymol. 308, 301–354. 31. Berti, P. J. (1999) Methods Enzymol. 308, 355–397. 32. Jiang, Y. L., and Stivers, J. T. (2001) Biochemistry 40, 7710 – 7719. 33. Sinnott, M. L. (1988) Adv. Phys. Org. Chem. 24, 113–204. 34. Shapiro, R., and Kang, S. (1969) Biochemistry 8, 1806 –1810. 35. Fujii, T., Saito, T., and Nakasaka, T. (1989) Chem. Pharmaceut. Bull. 37, 2601–2609. 36. Waters, T. R., and Swann, P. F. (1998) J. Biol. Chem. 273, 20007–20014. 37. Drohat, A. C., Jagadeesh, J., Ferguson, E., and Stivers, J. T. (1999) Biochemistry 38, 11866 –11875. 38. Noll, D. M., Gogos, A., Granek, J. A., and Clarke, N. D. (1999) Biochemistry 38, 6374 – 6379. 39. Riazuddin, S., and Lindahl, T. (1978) Biochemistry 17, 2110 – 2118. 40. Rachofsky, E. L., Seibert, E., Stivers, J. T., Osman, R., and Ross, J. B. Alexander (2001) Biochemistry 40, 957–967.
MINIREVIEW Uracil DNA Glycosylase: Insights from a Master Catalyst 1 James T. Stivers 2 and Alexander C. Drohat Department of Pharmacology and Molecular Sciences, The Johns Hopkins University School of Medicine, 725 North Wolfe Street, Baltimore, Maryland 21205-2185
Received September 12, 2001
The recognition and removal of damaged bases in the genome is the province of a highly specialized assemblage of enzymes known as DNA glycosylases. In recent years, structural and mechanistic studies have rapidly moved forward such that in some cases, the high-resolution structures of all stable complexes along the reaction pathway are available. In parallel, advances in isotopic labeling of DNA have allowed the determination of a transition state structure of a DNA repair glycosylase using kinetic isotope effect methods. The use of stable substrate analogs and fluorescent probes have provided methods for real time measurement of the critical step of damaged base flipping. Taken together, these synergistic structural and chemical approaches have elevated our understanding of DNA repair enzymology to the level previously attained in only a select few enzymatic systems. This review summarizes recent studies of the paradigm enzyme, uracil DNA glycosylase, and discusses future areas for investigation in this field. © 2001 Elsevier Science Key Words: DNA glycosylase; base flipping; inhibition; chemical rescue; transition state structure; enzyme mechanism.
DNA has evolved to be the genetic storage material because of its exceptional, but not infinite chemical stability. Nevertheless, genomic DNA is often viewed as a frail target that suffers the onslaught of a variety of chemical insults resulting in deamination, oxidation, alkylation, or other forms of chemical damage to the natural bases (1). Such damage, although infrequent, 1 The work in the author’s laboratory is supported by National Institutes of Health Grant GM46835. 2 To whom correspondence and reprint requests should be addressed. Fax: (410) 955-3023. E-mail: [email protected].
0003-9861/01 $35.00 © 2001 Elsevier Science All rights reserved.
is intolerable with respect to the preservation of the coding potential of the genome, and exerts the selection pressure for evolution of an elaborate cellular DNA repair machinery. Paradoxically, the intrinsic stability of DNA makes the breaking of its chemical bonds during DNA repair an energetically challenging feat, and therefore requires the action of powerful enzyme catalysts to efficiently recognize and remove damaged bases through a pathway known as base excision repair (Fig. 1). The first step in this repair pathway is the hydrolytic cleavage of the N-glycosidic bond of the damaged base by a damage specific DNA glycosylase, which is the focus of this review. DNA glycosylases face two fundamental obstacles in the initiation of this repair process. First, these enzymes must solve the formidable challenge of locating damaged bases in the vast genome before these potentially mutagenic sites are fixed by DNA replication. This critical time frame provides the driving force for selection of efficient damage search and detection mechanisms. Such mechanisms may involve three-dimensional or one-dimensional search processes (2), or even processive scanning mechanisms in which unusual features of the damage site are detected by the enzyme (3). A number of high resolution crystal structures of DNA repair glycosylases bound to DNA have appeared in recent years that have revealed several unifying principles of the damage recognition process (4 –10). All monofunctional DNA repair glycosylases studied to date operate by a base flipping mechanism in which the damaged base is extruded from the DNA base stack into the active site pocket of the enzyme, which is highly complementary to the shape and electronic features of the base. The remarkable conservation of the base flipping mechanism attests to the importance of gaining access to the functional groups on the base and sugar, which provides the binding inter1
2
STIVERS AND DROHAT TABLE I
Stabilities of Representative Deoxynucleosides at pH 7 and pH 3 and Estimated Catalytic Powers of Several DNA Glycosylases a
Nucleoside
t 1/2 (pH 7, 37°C)
t 1/2 (pH 3, 37°C)
DNA context
Specific DNA glycosylase
Catalytic power b (pH 7, 37°C)
T dU dA N 3-Me-dA
109 years 23 years 3.9 years 36 min
109 years 23 years 6.8 h 3 min
T/G U/A, U/G A/8-oxo-dG N 3-Me-dA/T
TDG UDG MutY TAG
7.5 ⫻ 10 7 2 ⫻ 10 12 4 ⫻ 10 7 ⱖ6
a This is not a comprehensive survey of all types of damaged bases, but does cover the entire range of stabilities that are found in nature. The half-lives for the nucleosides were calculated from the rate constants and activation parameters in references (20, 34, 35). b The catalytic power is defined as the half-life for the nonenzymatic reaction divided by that for the enzymatic reaction and is a measure of how much the enzyme lowers the activation barrier as compared to the nonenzymatic reaction. The enzymatic rates were obtained from single turnover measurements, such that slow product release rates do not affect the comparison. The enzymatic rates were obtained from the following studies: thymine DNA glycosylase (TDG) (36); uracil DNA glycoyslase (37); adenine mismatch-specific glycosylase (MutY) (38); and 3-methyladenine DNA glycosylase I from E. coli (TAG) (39).
actions necessary for lowering the activation barrier of the reaction. The stability of the glycosidic bond in pyrimidine, purine and alkylated purine deoxynucleosides varies greatly (Table I), which places differing catalytic demands on each of these enzymes. However, there are two fundamental chemical problems that each enzyme must solve to varying degrees. The first involves providing a mechanism for stabilization of the increased electron density that accumulates on the base during
bond cleavage. The second is balancing the degree of bond formation to the water nucleophile and bond breakage to the leaving group base in the transition state. The latter issue is a key mechanistic point because if the base departs completely before the water nucleophile attacks, the enzyme active site architecture must be designed to stabilize a highly unstable positively charged oxacarbenium ion intermediate (Fig. 1). In contrast, a transition state involving a large amount of bond formation to the water nucleophile
FIG. 1. Pathway for DNA base excision repair. Cellular DNA is damaged by a variety of exogenous and endogenous agents, resulting in alterations in the covalent structure of the natural bases (B*). The first step in repair is the hydrolytic cleavage of the N-glycosidic bond by a damage specific DNA glycosylase. The chemical mechanism of bond cleavage may be associative (upper pathway), or dissociative (lower pathway) involving a discrete positively charged oxacarbenium ion intermediate (see text). The repair process is completed by the action of an apyrimidinic endonuclease (AP endo), a repair polymerase (pol ), and DNA ligase that collectively remove the abasic nucleotide and reinsert the correct dNTP.
DNA GLYCOSYLASE MECHANISMS
3
requires activation of the anomeric carbon atom, which is weakly electrophilic, as well as the attacking water, which is a weak nucleophile. Of course, these issues are not new, and simply illustrate another example of the long-standing question of whether enzymes exert their power by stabilizing otherwise unstable intermediates (lower pathway, Fig. 1), or use catalytic strategies to lower the energy of concerted transition states (upper pathway, Fig. 1) (11). As will be discussed below, the electronic features of the base and the deoxyribose sugar in the transition state or high energy intermediate are likely to play a key role in the rational design of inhibitors of these enzymes. The intent of this review is to summarize some of the chemical approaches that have been used in recent years to elucidate the site recognition and catalytic mechanisms of the enzyme, uracil DNA glycosylase (UDG). 3 An emphasis is placed on UDG because the structural and mechanistic studies have advanced most rapidly in this system, and should therefore serve as a paradigm for similar studies of other glycosylases. DAMAGE RECOGNITION AND URACIL FLIPPING
Uracil in DNA may arise from either the spontaneous deamination of cytosine, resulting in U:G mismatches, or from the misincorporation of dUTP instead of TTP during DNA replication, which results in U:A base pairs (12). Such processes occur with alarming frequencies in normal cells (10 2 to 10 3 per cell per day), requiring constant surveillance and removal. The selective removal of a uracil base in the context of U:G or U:A base pairs, without also removing the structurally related base T, is one of the most difficult specific recognition problems in DNA repair. The entire process of damage site recognition is still not very well understood, but may be conceptualized as a three step process: (1) a three-dimensional or one dimensional search process in which the site is located in a sea of normal DNA base pairs, (2) the formation of largely nonspecific interactions with the site prior to base flipping, and (3) the formation of specific interactions with the flipped out nucleotide. The high-resolution crystal structures of UDG bound to specific DNA have contributed greatly to an understanding of the final step of extrahelical recognition (7, 10, 13). However, an understanding of the earlier steps requires either extrapolation backwards in time starting from the crystal models, or the use of other approaches that are better suited for understanding transient complexes. The specific recognition process of UDG is discussed within the context of this three-step model. 3 Abbreviations used: UDG, uracil DNA glycosylase; wt, wild-type; P, 2-aminopurine; MUG, mismatch uracil DNA glycoylase; TDG, thymine DNA glycosylase.
FIG. 2. (A) Structural model of the active site interactions of human UDG bound to duplex DNA containing the C-glycoside analogue 2⬘-deoxypseudouridine (dU) (PDB code 1 emh). Various residues proposed to be involved in chemical catalysis and glycosidic bond strain are indicted (see text); the residue numbering corresponds to the E. coli enzyme. (B) Hypothetical tautomerization reaction of dU that could account for the highly unusual distorted tetrahedral geometry of C1 observed in the crystal structure in A (see text).
Structure of the extrahelical complex. The crystal structure of human UDG bound to DNA containing the stable C-glycoside analog of deoxyuridine, deoxypseudouridine (dU, Fig. 2A, 2B), reveals the salient features of the site specific complex in which the uracil base is extrahelical. Aside from the unusual glycosidic bond angle seen in this structure (discussed later), this complex shares several features observed in other glycosylase complexes with extrahelical bases. First, UDG is found to use hydrogen bond donors (Ser88, Ser189) to clamp and pinch the phosphodiester backbone of the DNA on the same strand that contains the uracil base, leading to localized DNA bending (⬃45°). Another conserved aspect of base flipping seen in this structure is that a bulky hydrophobic amino acid side chain (Leu191, not shown for clarity) is found to project into the minor groove of the DNA duplex, filling the void left by the departed uracil base. The role of this enzyme group may be to actively push the damaged base from
4
STIVERS AND DROHAT
FIG. 3. Conceptual model for uracil base flipping by UDG. The overall process of base flipping may be viewed as being comprised of two components: an unfavorable duplex destabilization term (⌬G D), and a favorable stabilizing free energy (⌬G S) arising from all the interactions gained by flipping the base into the active site. Although poorly characterized, duplex destabilization is thought to arise in part from phosphodiester compression that occurs when UDG binds nonspecifically to DNA. The favorable free energy term arises in part from specific hydrogen bonding and stacking interactions with the uracil base, and intercalation of Leu191 into the DNA base stack to hinder reinsertion of the departed uracil.
the DNA stack, or alternatively, simply to hinder its reinsertion once the extrahelical state is attained by other means (see below). Finally, several completely conserved enzyme side chains form specific hydrogen bonds and pull the uracil base into the binding pocket: the O4 and H3 Watson-Crick acceptor and donor pair interact with the side chain atoms of Asn123, and uracil O2 forms hydrogen bonds to N 2H of His187, and the backbone amide of Gln63 (not shown). To venerate these observed interactions, a “pinch–push–pull” mechanism for base flipping has been proposed (6). Another key realization from this structure is that the specificity for uracil, as opposed to thymine, likely arises from unfavorable steric interactions of the 5-methyl group of T with Tyr66, which forms a rigid wall on one side of the binding pocket. Consistent with this hypothesis, removal of Tyr66 converts UDG into a thymine DNA glycosylase (14). Detecting transient complexes on the base flipping pathway. How do the initial interactions with the damage site and undamaged DNA lead to uracil flipping? Currently, there is no structure of a DNA glycosylase bound to undamaged DNA, or alternatively, to a damaged DNA site in which the base is not extrahelical. Thus, an entirely satisfying answer to this question is not yet in hand— but a trail for pursuit does exist. As suggested by both structural and biophysical studies, the earliest interaction of UDG with DNA probably involves local compression of the DNA phos-
phodiester backbone, perhaps leading to torsional stress in the DNA duplex. Upon encounter with the uracil site, the stress is presumably relieved by flipping the entire deoxyuridine nucleotide from the helical stack into the active site pocket of the enzyme (Fig. 3). The evidence for such a strain and release mechanism is indirect, but compelling. First, UDG has been shown to bind weakly to undamaged DNA (K D ⫽ 2 to 5 M), and much more tightly to DNA that contains the stable substrate analog 2⬘--fluoro-2⬘-deoxyuridine (2⬘FdU, K D ⬃25 nM). Raman spectroscopy and chemical probe methods have begun to characterize some features of the nonspecific and specific interactions with DNA. One spectral feature of the nonspecific complex was Raman hypochromism (analogous to UV hypochromism), arising from increased base stacking interactions in the DNA upon UDG binding (15). This spectral signature was also observed in the specific complex with substrate analog DNA, suggesting shared structural features in both complexes. Since Raman is a very sensitive probe for – interactions, the increase in stacking could be reporting indirectly on DNA distortion arising from enzyme interactions with the phosphodiester backbone. In a complementary approach, permanganate oxidation sensitivity experiments have been used to probe the change in accessibility of thymine bases upon binding of UDG to an undamaged 24-mer duplex DNA (Y. L. Jiang and J. T. Stivers, unpublished). These experiments clearly showed that UDG binding increased the oxidation sensitivity of specific T residues, to a level approaching that of the same sequence in a ssDNA context. These results lead us to hypothesize that UDG induces a subtle localized distortion in the duplex upon nonspecific binding that can destabilize base pairs some distance from the site of interaction. Although neither the Raman or chemical probe results can unambiguously establish that duplex destabilization is occurring, both methods reveal changes in the duplex structure that are evocative of such a mechanism. Another useful approach to study the steps leading to base flipping has been to use the fluorescent nucleotide probe 2-aminopurine (2-AP) in combination with the stable dU substrate analogue, 2⬘-fluoro-2⬘-deoxyuridine (2⬘-FdU) (16). 2-Aminopurine is an adenine analogue that has the useful properties of retaining base pair complementarity with T (although one hydrogen bond is moved to the minor groove of the duplex) and a fluorescence quantum yield that is very sensitive to stacking interactions with other bases in the duplex (17). Accordingly, when 2-AP ( ex ⫽ 315 nm, em ⫽ 370) is placed adjacent to 2⬘-FdU in duplex DNA, there is a large decrease in the stacking interaction when the uracil is flipped into the UDG active site, resulting in an eightfold increase in fluorescence. Such an approach provided the basis for a stopped-flow ki-
DNA GLYCOSYLASE MECHANISMS
netic analysis that revealed a two-step mechanism for damage site recognition and uracil flipping by UDG. In the first step, UDG forms an unstable complex in a diffusion controlled association reaction (K D ⬃ 2 M). It is noteworthy that this complex is kinetically and thermodynamically indistinguishable from the complex UDG forms with undamaged DNA, suggesting that the initial stage in site recognition is the formation of nonspecific interactions with the duplex. Once the weak complex forms, it may then decay backwards to form free reactants (k diss ⬃ 700 s ⫺1) or forward to form the specific extrahelical conformation (k flip ⬃ 1200 s ⫺1). The extremely rapid rate of dissociation from the weak complex is not consistent with a highly processive one-dimensional search mechanism, but is consistent with previous processivity measurements of eUDG (in the presence of physiological concentrations of salt), which indicated the enzyme did not linearly diffuse long distances on the DNA before dissociation (x). Another interesting finding from the fluorescence studies was that the internal equilibrium constant for base flipping on the enzyme (K flip ⫽ 12 to 33) was similar for duplex DNA containing 2⬘-FdU/A and 2⬘FdU/G base pairs, as well as single stranded 2⬘-FUcontaining DNA. This key observation provides additional support for a duplex destabilization step prior to base flipping, because it is difficult to rationalize the similar base flipping rates and equilibria for duplex and single stranded DNA unless the duplex DNA was destabilized prior to the flipping step. Stopped-flow tryptophan fluorescence measurements also indicate that an induced fit conformational change in UDG accompanies the process of uracil flipping. The conformational change was also detected by Raman spectroscopy when UDG binds to DNA containing 2⬘-FdU, but not undamaged DNA, indicating that the conformation of the enzyme is altered by the specific interactions that are formed as a result of base flipping (15, 16). Such a conformational change, which occurs concomitantly with base flipping, argues strongly for direct involvement of the enzyme in the formation of the extrahelical state, rather than passive trapping of a spontaneously formed extrahelical base (18). Manipulating base flipping using mutagenesis and chemical approaches. Two approaches to test our understanding of a biological reaction are to generate mutations, or alternatively, synthesize substrate analogs that remove functional groups that are hypothesized to play a role in the process. Conversely, the generation of substrate analogs or small molecules that “rescue” the damaging effects of enzyme mutations can provide valuable insights into the role of the enzyme functional group in the mechanism. One novel example of the chemical rescue approach was recently employed
5
FIG. 4. A pyrene nucleotide to promotes uracil flipping. (A) The bulky pyrene nucleotide fills the entire space usually occupied by a normal base pair. Thus substitution of pyrene (Y) for the A opposite to U (i.e., U/A 3 U/Y) would be expected to push the uracil from the base stack and facilitate specific DNA binding by diminishing the free energy penalty for duplex destabilization prior to uracil flipping (see Fig. 3). (B) Schematic depiction of the mechanism of pyrene (hashed symbol) assisted flipping (19).
to probe the pushing role of Leu191 in uracil flipping by UDG (19). Deletion of Leu191 results in a 10 to 625-fold decrease in k cat/K m using duplex DNA substrates containing a U:A base pair, presumably because UDG can no longer stabilize the extrahelical uracil in the active site. This interpretation was tested by substituting a pyrene nucleotide analogue for adenine opposite to the uracil (Fig. 4A). This rational design approach was based on the idea that the bulky pyrene “base,” which fills the entire space normally occupied by the normal U:A base pair (Fig. 4B), might serve as a mechanical wedge to either force the uracil from the DNA base stack in the free DNA (pushing) or hinder its reinsertion once it is expelled in the UDG complex (plugging). We predicted that Y would selectively rescue the L191A mutant of UDG that is defective in promoting the extrahelical base and would have a lesser effect on wtUDG and other mutants that were not defective in the base flipping step. Consistent with these expectations, pyrene opposite to U (a U:Y pair) selectively and completely rescued the impaired catalytic and binding activities of the L191A pushing mutant of UDG, with only a small salutary effect on mutants that were not deficient in base flipping. A model was proposed in which pyrene diminishes the energy penalty for flipping the base from the duplex, and stabilizes the extrahelical state. Pyrene rescue should serve as a useful tool to diagnose the functional roles of other residues involved in promoting the extrahelical base.
6
STIVERS AND DROHAT
CATALYTIC STRATEGIES
Glycosidic bond hydrolysis and the intrinsic chemical properties of deoxynucleotides. Before considering the nature of enzymatic catalysis it is always useful to identify the character of the chemical problem through consideration of the nonenzymatic reaction. The halflives and pH dependences of the nonenzymatic glycosidic bond cleavage reactions of pyrimidines, purines, and alkylated purine deoxynucleosides illustrate the point that the various DNA glycosylases specific for these base types face unique catalytic problems arising from the different intrinsic chemical properties of the bases (Table I). Pyrimidine-specific enzymes such as uracil DNA glycosylase (UDG) and the U:G and T:G mismatch glycosylases, MUG and TDG, face the most difficult leaving group activation problem because U and T cannot be readily protonated (pK a ⬍ ⫺3.4) (20). Purine bases are quite stable to hydrolysis at neutral pH (t 1/ 2 ⬃ 4 yrs). However, these bases are more easily protonated (pK aN1 ⫽ 3.5, pK aN7 ⬍ 1) (21), which provides a powerful mechanism for enzymatic activation of these bases by enzymes such as MutY (Table I). Finally, alkylated purine bases, and especially 3-MeA, are quite labile even at neutral pH (t 1/ 2 ⫽ 36 min), and may not require any activation at all by the alkyl purine specific enzymes such as 3-methyladenine DNA glycosylase I from Escherichia coli (TAG, Table I). Accordingly, the catalytic power of these enzymes 4 varies wildly from as little as 10-fold for TAG, to as much as 10 12-fold for UDG at pH 7 and 37°C. Of course, leaving group activation is not the only catalytic strategy that may be employed. If the enzymatic reaction pathway follows a highly dissociative mechanism in which a positively charged oxacarbenium ion is generated (Fig. 1), catalysis would be enhanced by any strategy that leads to stabilization of this cationic species. Such strategies could involve electrostatic mechanisms, or the use of binding energy to distort the sugar into a conformation that favors stabilizing hyperconjugative effects from the 2⬘ hydrogens in the dissociative transition state or intermediate (22). If an associative mechanism involving significant bond formation to the water nucleophile is followed, then such strategies as general base catalysis of water attack and polarization of the anomeric carbon would become useful. The following discussion of UDG uses the available structural and mechanistic information to evaluate the significance of these potential catalytic strategies. It is likely that related strategies will be used by other DNA glycosylases. 4 In this review, the catalytic power of an enzyme is defined as the single-turnover rate of the enzyme catalyzed reaction divided by the rate of the uncatalyzed reaction. Although k cat is often used for this comparison, this rate constant is less meaningful in the case of DNA glycosylases because the product release step is rate limiting.
Evaluating ground state strain and stereoelectronic effects with UDG. UDG is the most powerful DNA glycosylase yet identified. The enzyme provides a rate acceleration of 10 12-fold over the uncatalyzed hydrolysis reaction at 25°C, placing it in the upper echelon of enzyme catalysts. The study of UDG has been surprisingly amenable to a wide range of biophysical approaches. Extensive crystallographic studies on the human and E. coli enzymes have yielded structures of each complex along the reaction pathway (6, 7, 10, 23). In addition, heteronuclear NMR, Raman spectroscopic and kinetic isotope effect studies have provided unique perspectives into each reaction complex, including the transition state. One of the least understood and most interesting aspects of UDG catalysis is the role of ground state strain and stereoelectronic effects. These questions were propelled into serious consideration by the provocative and compelling crystal structure of human UDG in complex with DNA containing the stable Cglycoside substrate analog 2⬘-deoxypseudouridine (⌿dU, Fig. 2A). This structure indicates that the normal trigonal planar C1 position of ⌿dU is bent toward a tetrahedral geometry, presumably through steric interactions with a conserved phenylalanine in the active site, and the 3⬘ and 5⬘ serine phosphodiester clamps. It was argued that such an unusual conformation, if affected in the natural substrate, would lead to groundstate bond strain and enhanced electron orbital overlap between the lone pair electrons on O4⬘ and the -orbitals of the uracil ring in the transition state, thereby solving a problem described as the “orthogonal lone pair paradox” (7). 5 The interpretation of this structure requires contemplation of two separate questions. First, is ⌿dU-DNA a faithful substrate mimic? And second, is such a destabilization mechanism energetically reasonable and, more importantly, catalytic? The available evidence indicates that ⌿dU is not the best available substrate mimic for UDG. DNA containing ⌿dU binds 20-fold more weakly than the same DNA sequence containing the alternative slow substrate and competitive inhibitor 2⬘-FdU (Y. L. Jiang and J. T. Stivers, unpublished), which has a K D similar to the K m of the natural 2⬘-dU-containing substrate (16). It is also known from Raman spectroscopy studies that 2⬘-FdU in DNA shows a large 34 cm ⫺1 downshift in 5
The term paradox was used to describe the orthogonal arrangement of the * orbital of the anomeric carbon and the orbital of the C2–O2 bond of the uracil ring in the normal anti-conformation of deoxyuridine (see Fig. 5 of Ref. 7). There is a long standing debate on the energetic importance of such stereoelectronic effects (33). In the present case, there is experimental evidence from Raman spectroscopy for N ␦⫹ ⫽ C-O ␦⫺ zwitterionic character at N1–C2 ⫽ O in the ground state. The presence of such a resonance form suggests that alignment of orbitals would not be required for efficient leaving group expulsion in the transition state.
DNA GLYCOSYLASE MECHANISMS
the carbonyl stretching frequencies of the uracil ring upon binding to UDG, indicating strong hydrogen bonding to active site groups (Fig. 2A), whereas ⌿dU containing DNA does not show any significant polarization of these carbonyl groups (J. Dong and P. R. Carey, unpublished). An additional concern, that is entirely unresolved, is that the observed structure for ⌿dU in the crystal could arise not from mechanical strain, but from a tautomerization reaction, in which the H3 proton is transferred to C1 (Fig. 2B). Indeed, the observed crystal conformation and the computationally optimized structure of the C1-H tautomer of deoxypseudouridine are very similar, and probably beyond the resolution of the diffraction data (1.8 Å). Although QM calculations indicate that this tautomerization reaction is thermodynamically unfavorable in solution by about 20 kcal/ mol, due to the loss of aromaticity in the ring (J. Dong and P. R. Carey, unpublished), the active site environment of UDG could kinetically facilitate this reaction through the use of the strong H-bond with His187 (Fig. 2, see also below). The proton could become trapped at C1 if no basic group is available to accept the proton in the tetrahedral tautomer (no such group is seen in the crystal structure). Although this thermodynamic barrier is large, there are many enzymes in Nature that catalyze similar unfavorable tautomerization reactions using electrophilic mechanisms (24). A perplexing observation is that no unusual changes in the normal modes of the uracil ring were observed in solution Raman studies of 2⬘-FdU bound to UDG, as would be expected from such bond distortion. This is intriguing because 2⬘-FdU cannot undergo the same tautomerization reaction as ⌿dU. In summary, the questions raised by this structure have large implications for the role of strain in enzyme catalysis, and more specifically, in the design of tight binding inhibitors of UDG. A rational viewpoint for the enzymologist at this juncture is to await the results of further structural or spectroscopic studies that will undoubtedly clarify the role of ground state strain in this system. If such a mechanism is found to prevail, it will be fascinating to dissect the mechanical forces that give rise to such a remarkable effect. Nature of the transition state. Significant advances have been made in understanding the electronic and structural features of the transition state of the UDG reaction using spectroscopic and kinetic isotope effect methodologies. The first hint at the electronic character of the transition state came from 2D 13C heteronuclear NMR studies, in which the ionization state of the uracil base in the ternary product complex with abasic DNA was determined (25). Quite surprisingly, the uracil was found to have a dramatically lowered N1 pK a, such that the uracil anion was the sole bound
7
species at neutral pH (pK aN1 ⫽ 6.4). The presence of the uracil anion was later confirmed in Raman studies (15). Taken together, these observations indicated that the enzyme stabilized the uracil anion leaving group by 5 kcal/mol in the product complex as compared to free uracil in solution (pK aN1 ⫽ 9.8)— but how? NMR spectroscopy also illuminated the answer to this energetic problem. Inspection of the 1D proton NMR spectrum of the UDG complex with uracil and abasic DNA revealed a downfield shifted proton (␦ ⫽ 15.7 ppm), which was unambiguously assigned to a short (⬍2.7 Å) hydrogen bond between the O2 anion of uracil and the N 2 H of the conserved His187 (26). Histidine 187 was previously shown by NMR to be neutral in the pH range 4.5–10, which, in addition to the other NMR evidence, established the neutral histidine-uracil anion interaction depicted in Fig. 5. This provided yet another example in which evolution has selected for a short strong hydrogen bond from a neutral histidine to stabilize a developing negative charge on an enolic oxygen atom (27). Thus, part of the catalytic story of UDG involves leaving group stabilization by hydrogen bonding, but not proton transfer. An inference from these NMR studies was that the uracil anion might also exist as an intermediate in a stepwise mechanism for glycosidic bond cleavage. The direct determination of the transition state structure for the UDG catalyzed reaction was obtained from kinetic isotope effect measurements. This approach required the development of methods for the enzymatic synthesis of oligonucleotide substrates containing stable and radioactive isotope labels at a single uracil residue (28, 29). As discussed in two recent and excellent reviews by Schramm and Berti (30, 31), kinetic isotope effects (KIE ⫽ k light/k heavy) measure the changes in vibrational frequencies for heavy and light isotopes incorporated at specific atomic sites of a reactant as it is transformed from its reactant state to the transition state geometry, and thus provide information on bond lengths and geometries in the transition state. The experimental KIEs for UDG unambiguously indicated a dissociative transition state with substantial oxacarbenium ion character, and confirmed the proposal that a short-lived oxacarbenium ion-uracil anion intermediate is formed on the enzyme (Fig. 5). One remarkable feature of this intermediate was that the sugar adopted an unusual 3⬘-exo pucker, which allows maximal stabilizing hyperconjugative effects from the 2⬘  protons to the electron deficient anomeric carbon. This sugar conformation in the intermediate was strikingly similar to that observed in the crystal structure of the Michaelis complex, suggesting that UDG uses binding energy in the ground state to preorganize the substrate into this reactive conformation. We concluded that several features of the active site contributed to the increased stability of the intermediate as
8
STIVERS AND DROHAT
FIG. 5. Interactions leading to stabilization of the oxacarbenium ion-uracil anion intermediate in the UDG active site. The electrostatic potentials for the dU reactant and the oxacarbenium ion intermediate are projected onto the van der Waals surfaces for each molecule. The structures were derived using the geometric information derived from KIE measurements (28), and the crystal coordinates of the product complex with the uracil anion and abasic DNA (PDB code 1 ssp). For clarity, His187 is not shown in the depiction of the electrostatic potentials. Stabilization of the ionic intermediate is thought to arise from the enforcement of stabilizing hyperconjugative effects in the reactant and transition states using the serine phosphodiester interactions, and (ii) electrostatic stabilization of the oxacarbenium ion by the anionic uracil leaving group and Asp64.
compared to solution: (i) the enforcement of stabilizing hyperconjugative effects in the reactant and transition states using the serine phosphodiester interactions, and (ii) electrostatic stabilization of the oxacarbenium ion by the anionic uracil leaving group and Asp64 (Fig. 5). Substrate binding energy and catalysis. Although enzymes are sometimes viewed as elaborate scaffolds for carrying chemical catalysts, the intricate interplay between noncovalent interactions with the substrate that are distant from the site of chemistry can play an important role in catalysis. Such interactions are critical for obtaining proper substrate positioning with respect to catalytic groups, driving conformational changes in the substrate that enhance its reactivity, and in induced fit specificity mechanisms. One approach to investigate these effects with UDG was to systematically dismantle the optimal substrate, Ap ⫹1UpA ⫺1pA ⫺2, by replacing the nucleotides at the ⫹1, ⫺1, or ⫺2 positions with a tetrahydrofuran abasic site analogue, 3-hydroxypropyl phosphodiester spacer, a phosphate monoester, or a hydroxyl group (32). Thus, base, phosphodiester, and deoxyribose interactions were removed in a controlled fashion, and the damaging effects on catalysis could be ascertained. One key finding in this extensive study was that the minimal substrate for UDG is deoxyuridine (dU). Although four billion-fold less reactive than the optimal substrate, cleavage of the glycosidic bond of dU was still enhanced by 4 ⫻ 10 7-fold over the spontaneous reaction in the absence of UDG. A second significant finding was the tremendous importance of the ⫺2 nucleotide in driving
optimal positioning of the substrate. The net catalytic benefit of adding a ⫺2 adenine nucleotide to ApUpA (i.e., ApUpA 3 ApUpApA) was 270,000-fold (⫺7.5 kcal/ mol). The crystal structure suggests a structural basis for this large effect because the pro-Sp nonbridging oxygen atom of the ⫺2 phosphodiester is hydrogen bonded to the backbone amide group of the catalytic electrophile His187, which is in the same loop as Leu191 (7). Thus the ⫺2 nucleotide likely serves as a key handle to drive the active site into the productive closed conformation, allowing formation of the strong hydrogen bond between uracil O2 and His187, optimizing the serine-phosphodiester interactions at the ⫹1 and ⫺1 positions, and positioning Leu191 in the minor groove. FUTURE PROSPECTS
There still remain many interesting questions to be investigated with respect to the mechanism of damage site recognition and base flipping by UDG and other repair glycosylases. One exciting area for exploration would be the use of single molecule fluorescence methods to visualize the process of damage site location one molecule at a time. New approaches also need to be used to understand the structure and dynamics of the nonspecific complex between UDG and undamaged DNA. It is likely that NMR will find great utility in this respect, as it may be possible to determine the structure of duplex DNA in such a complex, and to investigate the dynamics by measuring imino proton exchange rates or performing 13C or 15N relaxation mea-
DNA GLYCOSYLASE MECHANISMS
surements. These NMR dynamic studies could be augmented with time-resolved fluorescence measurements of the dynamic behavior of damaged DNA and its complex with the enzyme. Such investigations have already been performed on the complex of UDG with the abasic product, revealing that the enzyme restricts the conformational dynamics of the duplex compared to the free DNA (40). Finally, it would be of great utility to develop selective small molecule inhibitors of DNA glycosylases and other enzymes involved in DNA base excision repair. Cell permeable inhibitors could be used to better probe the biological role of this pathway under various stress conditions such as those used in cancer chemotherapy treatment. The pursuit of these and other goals will continue to make DNA repair an exciting field into the foreseeable future. REFERENCES 1. Friedberg, E. C. (1985) in DNA Repair, W. H. Freeman, New York, NY. 2. Berg, O. G., Winter, R. B., and von Hippel, P. H. (1981) Biochemistry 20, 6929 – 6948. 3. Verdine, G. L., and Bruner, S. D. (1997) Chem. Biol. 4, 329 –334. 4. Lau, A. Y., Scharer, O. D., Samson, L., Verdine, G. L., and Ellenberger, T. (1998) Cell 95, 249 –258. 5. Lau, A. Y., Wyatt, M. D., Glassner, B. J., Samson, L. D., and Ellenberger, T. (2000) Proc. Natl. Acad. Sci. USA 97, 13573– 13578. 6. Parikh, S. S., Mol, C. D., Slupphaug, G., Bharati, S., Krokan, H. E., and Tainer, J. A. (1998) EMBO. J. 17, 5214 –5226. 7. Parikh, S. S., Walcher, G., Jones, G. D., Slupphaug, G., Krokan, H. E., Blackburn, G. M., and Tainer, J. A. (2000) Proc. Natl. Acad. Sci. USA 97, 5083–5088. 8. Bruner, S. D., Norman, D. P., and Verdine, G. L. (2000) Nature 403, 859 – 866. 9. Barrett, T. E., Scharer, O. D., Savva, R., Brown, T., Jiricny, J., Verdine, G. L., and Pearl, L. H. (1999) EMBO J. 18, 6599 – 6609. 10. Werner, R. M., Jiang, Y. L., Gordley, R. G., Jagadeesh, G. J., Ladner, J. E., Xiao, G., Tordova, M., Gilliland, G. L., and Stivers, J. T. (2000) Biochemistry 39, 12585–12594. 11. Jencks, W. P. (1980) Accounts Chem. Res. 13, 161–169. 12. Mosbaugh, D. W., and Bennett, S. E. (1994) Prog. Nucleic Acid Res. Mol. Biol. 48, 315–370. 13. Parikh, S. S., Mol, C. D., and Tainer, J. A. (1997) Structure 5, 1543–1550.
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